Short title: Role of plasma membrane aquaporins in maize
1 2
Author for Contact details: François Chaumont, francois.chaumont@uclouvain.be
3 4 5
Modification of the expression of the aquaporin ZmPIP2;5 affects water
6
relations and plant growth
7
8 9
Lei Ding,a,1 Thomas Milhiet,a,1, Valentin Couvreur,b Hilde Nelissen,c,d Adel Meziane,a,e Boris
10
Parent,e Stijn Aesaert,c,d Mieke Van Lijsebettens,c,d Dirk Inzé,c,d François Tardieu,e Xavier
11
Draye,b François Chaumonta,2
12 13
a Louvain Institute of Biomolecular Science and Technology, UCLouvain, 1348
14
Louvain-la-Neuve, Belgium
15 16
b Earth and LifeInstitute, UCLouvain, 1348 Louvain-la-Neuve, Belgium
17 18
c Center for Plant Systems Biology, VIB-Ghent University, Technologiepark 71, 9052 Ghent,
19
Belgium
20 21
d Department of Plant Biotechnology and Bioinformatics, Ghent University, 9052 Gent,
22
Belgium
23 24
e Laboratoire d'Ecophysiologie des Plantes sous Stress Environnementaux (LEPSE), Université
25
de Montpellier, Institut National de la Recherche Agronomique (INRA), F-34000 Montpellier,
26 27 28
1These authors contributed equally to the article.
29
2Author for contact
30 31
One-sentence summary: Reverse genetic approaches demonstrate that the maize plasma
32
membrane PIP2;5 aquaporin plays a role in controlling root radial water movement, leaf
33
hydraulic conductivity, and plant growth.
34 35 36
Author contributions: L.D., T.M., H.N., B.P., and F.C. designed the experiments. L.D., T.M.,
37
H.N., A.M., B.P., and S.A. performed the experiments. M.V.L. supervised the maize
38
transformation. L.D., T.M., V.C., H.N., A.M., B.P., S.A., F.T., X.D., M.V.L., and F.C. analyzed
39
the data. L.D. and F.C. wrote the manuscript. All the authors contributed to the discussion and
40
revision of the manuscript.
41
Plant Physiology Preview. Published on January 24, 2020, as DOI:10.1104/pp.19.01183
42 43
Funding information: This work was supported by the Belgian National Fund for Scientific
44
Research (FNRS, FRFC 2.4.501.06F), the Interuniversity Attraction Poles Programme-Belgian
45
Science Policy (grant IAP7/29), the “Communauté française de Belgique-Actions de
46
Recherches Concertées” (grants ARC11/16-036 and ARC16/21-075) and the Pierre and Colette
47
Bauchau Award. L.D. was supported by an Incoming Post-doc Move-in Louvain Fellowships
48
co-funded by the Marie Curie Actions. T.M. was supported by a research fellow at the Fonds de
49
Formation à la Recherche dans l’Industrie et l’Agriculture.V.C. was supported by the FNRS
50
(FC 84104).
51 52
Abstract
53 54
In maize (Zea mays), the plasma membrane intrinsic protein PIP2;5 is the most highly
55
expressed aquaporin in roots. Here, we investigated how deregulation of PIP2;5 expression
56
affects water relations and growth using maize overexpressing (OE; B104 inbred) or knockout
57
(KO; W22 inbred) lines. The hydraulic conductivity of the cortex cells of roots grown
58
hydroponically was higher and lower in PIP2;5 OE and pip2;5 KO lines, respectively,
59
compared with their corresponding wild-type (WT) plants. While whole root conductivity
60
decreased in the KO lines compared to the WT, no difference was observed in OE plants. This
61
paradox was interpreted using the MECHA hydraulic model, which computes the radial flow
62
of water within root sections. The model hints that the plasma membrane permeability of the
63
cells is not radially uniform but PIP2;5 may be saturated in cell layers with apoplastic barriers,
64
i.e. the endodermis and exodermis, suggesting the presence of post-translational mechanisms
65
controlling the abundance of PIP in the plasma membrane in these cells. At the leaf level,
66
where the PIP2;5 gene is lowly expressed in WT plants, the hydraulic conductance was
67
higher in the PIP2;5 OE lines compared with the WT plants, whereas no difference was
68
observed in the pip2;5 KO lines. The temporal trend of leaf elongation rate used as a proxy of
69
that of xylem water potential was faster in PIP2;5 OE plants upon mild stress, but not in
70
well-watered condition, demonstrating that PIP2;5 may play a beneficial role for plant growth
71
under specific conditions.
72 73 74
Introduction
75 76
Aquaporins, belonging to the plasma membrane intrinsic protein (PIP) subfamily, are
77
major actors controlling membrane water permeability. The physiological functions of PIPs
78
are straightforward at the cell level (Hachez et al., 2006; Hachez et al., 2008; Hachez et al.,
79
2012; Heinen et al., 2014), but are considerably more complex at the organ and whole-plant
80
levels. For example, overexpression or silencing of PIP aquaporins has contrasting effects on
81
root hydraulics (Siefritz et al., 2002; Hachez et al., 2006; Postaire et al., 2010; Sade et al.,
82
2010), root development (Péret et al., 2012), leaf hydraulic conductance (Kleaf) (Prado and
83
Maurel, 2013; Sade et al., 2014), stomatal movement (Grondin et al., 2015; Wang et al., 2016;
84
Rodrigues et al., 2017), and transpiration (Tr) (Maurel et al., 2016). This is largely because the
85
transcellular path, in which water crosses the cell membranes mainly through aquaporins
86
(Steudle and Peterson, 1998; Steudle, 2000) occurs simultaneously together with other two
87
pathways, namely the apoplastic path, in which water flow goes through the cell wall, and the
88
symplastic path, in which water moves through the plasmodesmata. The overall root hydraulic
89
conductivity (Lpr) is the integration of conductivity from the three pathways; their
90
contribution to the Lpr varies according to root anatomy development and environmental
91
factors (drought, high salinity, nutrient availability, and anoxia, etc.). To better understand the
92
complexity of root radial hydraulic conductivity and integrate the multiple variables,
93
mathematical models have been developed (Steudle and Peterson, 1998; Zwieniecki et al.,
94
2002; Foster and Miklavcic, 2017; Couvreur et al., 2018). Among them, the ‘MECHA’ model
95
(Couvreur et al., 2018) predicts the root radial hydraulic conductivity, based on the detailed
96
radial anatomy of the root and the distribution of the cell wall hydraulic conductivity, the cell
97
plasma membrane permeability, the hydraulic conductance, the frequency of plasmodesmata,
98
and the membrane reflection coefficients. MECHA is therefore appropriate to address
99
subcellular hydraulics and its impact on the radial transport of water.
100
Here, we analyzed the effects of the overexpression or silencing of the maize (Zea mays)
101
PIP2;5 at both cellular and whole-plant levels. The PIP2;5 gene encodes an active water
102
channel (Fetter et al., 2004), is the most expressed PIP gene in the primary root (Hachez et al.,
103
2006), and shows a polarized localization at the plasma membrane side facing the external
104
medium, supporting a function in root water uptake. Another clue for the involvement of
105
PIP2;5 in radial water movement is its high expression in cell types with Casparian strips
106
(exodermis and endodermis), places where water has to enter the symplast to continue its flow
107
to the xylem vessels (Hachez et al., 2012). In roots, the expression of PIP2;5 mRNA and its
108
protein abundance are modulated by diurnal and circadian rhythm, osmotic stress, and growth
109
conditions (aeroponic and hydroponic) (Hachez et al., 2012; Caldeira et al., 2014). In addition,
110
PIP2;5 proteins are more or less abundant in maize lines overexpressing or silenced for an
111
ABA biosynthesis gene, respectively (Parent et al., 2009). The PIP2;5 gene is weakly
112
expressed in leaves, with a maximum of expression at the end of the elongation zone and the
113
zone where the leaf emerges from the sheath and where lignification of the metaxylem is
114
observed (Hachez et al., 2008). Altogether, these data suggest that PIP2;5 plays important
115
roles in regulating water relations in maize, but genetic approaches to further understand its
116
physiological function are missing.
117
We first characterized PIP2;5 overexpressing (OE) and pip2;5 knockout (KO) lines at the
118
molecular and cellular levels and addressed the question of the effects of the manipulation of
119
PIP2;5 expression at the plant level by collectively examining the hydraulic conductance in
120
roots and leaves and the time course of leaf elongation rate, considered here as a way to
121
indirectly assess the changes in xylem water potential, following fluctuations of the
122
evaporative demand (Caldeira et al., 2014; Caldeira et al., 2014). While the cell hydraulic
123
conductivity was affected by the deregulation of PIP2;5 expression, the contrasting results at
124
the organ levels suggest that upscaling requires a modeling approach to decipher the dataset
125
presented here.
126 127
Results
128
Generation, Isolation, and Molecular Characterization of Maize Lines Deregulated in
129
PIP2;5 Expression
130
To determine the role of PIP2;5 aquaporin in maize, we first prepared a genetic construct
131
aiming at constitutively overexpressing the PIP2;5 gene under the control of the p35S
132
promoter (Fig. 1A), and performed an Agrobacterium-mediated transformation of the inbred
133
B104. Two independent PIP2;5 overexpressing lines (PIP2;5 OE-4 and OE-13) with high
134
PIP2;5 protein content from Western blotting analysis (see below) were selected for further
135
molecular characterization. In addition, we obtained from the “Maize Genetic COOP Center”
136
(http://maizecoop.cropsci.uiuc.edu/) a putative pip2;5 knockout (pip2;5 KO) W22 inbred line
137
(UFMu00767) containing a Mu transposon in the PIP2;5 gene (Fig. 1B). The presence and
138
the site of insertion of the Mu transposon in PIP2;5 gene were determined by
139
PCR-amplification of genomic DNA using PIP2;5 and Mu specific primers; this showed that
140
the Mu transposon was inserted in the second intron of the PIP2;5 gene (Fig. 1B).
141
To confirm the overexpression and downregulation of PIP2;5 in the maize lines,
142
microsomal fractions were prepared from roots and leaves, and PIP2;5 was immunodetected
143
using specific antibodies (Hachez et al., 2006). In roots, where the endogenous PIP2;5 gene is
144
the highest expressed PIP gene, a 17% and 141% increase in PIP2;5 protein abundance in the
145
PIP2;5 OE-4 and OE-13 lines, respectively, were observed when compared with the
146
non-transgenic segregating siblings (B104, named afterwards WT-B104) (Fig. 1C). In leaves,
147
where the endogenous PIP2;5 is lowly expressed, more than 10-fold higher PIP2;5 protein
148
abundance was detected in the PIP2;5 OE-4 and OE-13 lines compared with the WT-B104
149
plants (Fig. 1D). On the other hand, the PIP2;5 protein level was ninefold lower in the roots of
150
pip2;5 KO line than in roots of non-transgenic segregating siblings (W22, named afterwards
151
WT-W22) (Fig. 1C). No significant difference in PIP2;5 signal intensity was found between
152
pip2;5 KO and WT-W22 leaves (Fig. 1D), but the PIP2;5 signals in leaves were hardly
153
detectable and only observed after a very long exposure.
154
We also analyzed the PIP2;5 mRNA levels in roots and leaves by RT-qPCR. In the OE
155
plants, while no difference in PIP2;5 endogenous mRNA levels were observed, a high PIP2;5
156
transgene mRNA signal was detected using a PIP2;5-specific forward primer and a construct
157
linker-specific reverse primer (Supplemental Fig. S1, A-D). No amplification was detected
158
with this pair of primers in the WT-B104 plants. In pip2;5 KO plants, we observed 2175-fold
159
and fivefold lower mRNA level in roots (Supplemental Fig. S1E) and leaves (Supplemental
160
Fig. S1F), respectively, than in the WT-W22 plants. To investigate the reason why a very faint
161
protein signal was still observed in the leaf extract by immunodetection, we performed
162
RT-qPCR with primers flanking the Mu insertion site and detected a weak signal
163
(Supplemental Fig. S2), suggesting that the pip2;5 KO plants are not a complete
164
loss-of-function line.
165
To investigate whether the expression of other PIPs was affected by the deregulation of
166
PIP2;5 gene in both roots and leaves, RT-qPCR and Western blotting were performed. No
167
significant difference in the mRNA levels of most PIPs was observed between PIP2;5 OE
168
lines or pip2;5 KO and their corresponding WT lines in both roots and leaves (Supplemental
169
Fig. S1). Similarly, at the protein level, no significant difference in PIP1;2 and PIP2;1/2;2 was
170
observed between the OE or KO lines and their respective WT plants (Supplemental Fig. S3).
171
Because we used two different maize genetic backgrounds in this work, we checked that
172
the expression pattern of PIP2;5 was similar in the W22 and B73 lines (Hachez et al., 2006,
173
2008 and 2012). Similar to the previous results obtained in the B73, an intense signal of
174
PIP2;5 was immunodetected in the exo- and endodermis cells of W22 root, where the lignin
175
and suberin are deposited (Supplemental Fig. S4, A and B). PIP2;5 was also the most highly
176
expressed PIP gene in W22 roots and was lowly expressed in leaves (Supplemental Fig. S4C).
177 178
Altered Hydraulic Conductance in the PIP2;5 Deregulated Plants
179
We first investigated the effect of PIP2;5 deregulation on the hydraulic conductivity of the
180
root cortex cell (Lpc), measured with a cell pressure probe. The half-time of water exchange
181
(T1/2) across the membrane of root cortex cells was 1.9 to 1.8 times shorter (faster water flow)
182
in the two PIP2;5 OE lines than in WT-B104, whereas T1/2 was 2.8 times longer (slower water
183
flow) in the pip2;5 KO line with respect to the WT-W22 (Table 1). As a result, the Lpc was 69%
184
and 67% higher in PIP2;5 OE-4 and OE-13 lines, respectively (Fig. 2, A and B). In contrast,
185
the Lpc decreased by 63% in the pip2;5 KO line compared with the WT-W22 (Fig. 2C). The
186
turgor pressure and the cell elastic modulus (ε) were not affected by the deregulation of
187
PIP2;5 expression (Table 1), with the exception of a higher εcorrected in PIP2;5 OE-13 lines than
188
in WT-B104 plants. Besides, a bigger cell volume was also measured in PIP2;5 OE-13 lines
189
than in WT-B104 plants, suggesting that PIP2;5 overexpression in this line has affected the
190
root cell expansion. Consistently, the membrane water permeability of leaf mesophyll
191
protoplasts (Pos)was significantly higher in both PIP2;5 OE lines than in WT-B104 plants (Fig.
192
2, D and E). In comparison with WT-B104, an 85% and 60% increase in Pos was observed in
193
PIP2;5 OE-4 and PIP2;5 OE-13 lines, respectively. On the other hand, no difference in Pos
194
was observed between the WT-W22 and pip2;5 KO lines (Fig. 2F), due to the fact that PIP2;5
195
is barely expressed in WT leaves. It is worth mentioning that the Lpc and Pos mean values were
196
higher in WT-W22 than in WT-B104, suggesting that both inbred lines have different intrinsic
197
membrane permeabilities.
198
Consistent with the results at the cell level, the leaf hydraulic conductance (Kleaf)
199
measured with a hydraulic conductance flow meter (HCFM), increased by 58 % and 171 % in
200
the PIP2;5 OE-4 and OE-13 lines, respectively, when compared with the Kleaf of WT-B104
201
plants (Fig. 3, D and E), whereas no significant difference in Kleaf was found between pip2;5
202
KO plants and WT-W22 plants (Fig. 3F). However, at the root level, the increase in Lpc did
203
not result in a significant difference in the whole root conductance (Lpr) between the
204
WT-B104 and the PIP2;5 OE lines (Fig. 3, A and B). Conversely, Lpr was significantly lower
205
in the pip2;5 KO line than in the WT-W22 (Fig. 3C).
206
The lack of correlation between cortex Lpc and Lpr in PIP2;5 OE plants is not a
207
straightforward result to decipher. There are two reasons for this: (1) There are multiple
208
hydraulic media in series across the root radius (including cell walls, membranes, and
209
plasmodesmata, whose hydraulic conductivity may vary in each cell layer). Lpr is mostly
210
sensitive to the hydraulic conductivity of media that limit water flow the most (e.g. at
211
gatekeeper cell layers, i.e. endodermis and exodermis, where water flow through cell walls is
212
limited by apoplastic barriers). (2) Root hydraulic media are arranged both in series and
213
parallel, so that water pathways may bypass some of the most limiting media (e.g. at
214
gatekeeper cells, water may bypass cell walls by flowing through membranes, and a fraction
215
of water flow also bypasses gatekeeper cell membranes by using a symplastic path). Hence,
216
the quantitative modeling tool MECHA (Couvreur et al., 2018) with subcellular resolution of
217
water flow was needed to statistically validate the significance of hypotheses possibly
218
explaining the lack of correlation between cortex Lpc and Lpr between WT and PIP2;5 OE
219
plants. Hypothesis A considered that plasma membrane Lpc is uniform across cell layers (Fig.
220
4A), while hypothesis B considered that PIP2;5 is already “saturated” in WT gatekeeper cells
221
(their Lpc equals that measured in the cortex of PIP2;5 OE lines) and “unsaturated” in other
222
cell layers (their Lpc equals that measured in the WT cortex) (Fig. 4E). Three values of cell
223
wall hydraulic conductivity (kw 1-3), spanning a range from the literature, were considered for
224
each hypothesis. Statistical analysis of results from this combined modeling and experimental
225
approach suggested that, given the observed Lpc in WT-B104 and PIP2;5 OE plants, radially
226
uniform patterns of cell membrane permeability may not account for the observed contrast
227
between Lpr in WT-B104 and PIP2;5 OE-4 line, regardless of the cell wall hydraulic
228
conductivity kw (Fig. 4C, p < 0.01). The simulations reproduced the observed contrasts
229
between PIP2;5 OE and KO lines (Fig. 4, F-H) at the conditions that kw was higher than
230
6.9×10-10 m2.s-1.MPa-1 (kw2), and that the contribution of PIP2;5 to Lpc was saturated in the
231
endodermis and exodermis of WT lines (Fig. 4E, p < 0.05).
232 233
Altered Plant Growth in PIP2;5 Deregulated Plants Under Water Deficit Conditions
234
To further investigate the effect of PIP2;5 deregulation on water relations and plant growth,
235
we examined the time course of leaf elongation rate (LER) over changes in environmental
236
conditions. During the day, while the evaporative demand increases, hydraulic resistances to
237
water transfer can rapidly decrease the leaf water potential (Ψleaf) and the leaf growth, but it is
238
only observable under suboptimal water conditions (Bouchabké et al., 2006). We therefore
239
measured the leaf water potential and expansion of PIP2;5 OE-4 plants and their WT-B104
240
under well-watered conditions (Fig. 5C, in order to observe putative intrinsic differences of
241
leaf expansion rate) and under moderate water deficit (Fig. 5D, in order to observe the effects
242
of hydraulics).
243
Under moderate water deficit (Ψsoil = -0.23 to -0.26 MPa), we measured a higher leaf
244
water potential in the PIP2;5 OE-4 line than in WT-B104 plants, while the difference was not
245
observed under well-watered conditions (Fig. 5B). A significantly faster recovery of LER was
246
observed after the early-morning drop in PIP2;5 OE-4 compared with WT-B104 plants
247
resulting in large differences of LER during the day (Fig. 5D, inset). Overall, the mean LER
248
of PIP2;5 was higher in OE-4 than in WT-B104 over one day (Supplemental Fig. S5, 0.75
249
mm.h-1 vs 0.40 mm.h-1). During the night, the average LER of PIP2;5 OE-4 was 1.42 mm.h-1
250
and it was also significantly higher (p< 0.001) than the average LER of WT-B104 (1.05
251
mm.h-1). As expected, these differences were not observed in well-watered conditions (Fig.
252
5C), indicating that no pleiotropic effects had affected the intrinsic potential leaf expansion
253
rate of OE plants.
254
A faster recovery of LER was correlated with an increase in PIP aquaporin expression,
255
Ψleaf and Lpr (Parent et al., 2009; Caldeira et al., 2014). While no change in Lpr was recorded
256
in PIP2;5 OE plants compared with the WT-B104 plants in well-watered condition, we
257
compared the Lpr in response to short term osmotic stress (10% w/v polyethylene glycol (PEG)
258
6000; Ψ = -0.15 MPa) and observed a higher Lpr, but not significantly different, in PIP2;5
259
OE-4 than in WT-B104 plants (Supplemental Fig. S6). These results suggest that the higher
260
root hydraulic conductivity for PIP2;5 OE-4 observed under low osmotic potential translated
261
into differences of leaf water potential and leaf expansion under moderate water deficit, which
262
was not the case under well-watered conditions due to the very low difference of root
263
hydraulic conductivity.
264 265
Discussion
266 267
A better understanding of the functional role of PIP aquaporins in plant water relations is
268
essential to develop crop lines that use water more efficiently and are more tolerant to water
269
deficit. To this aim, we investigated the direct contribution of PIP2;5, the most expressed PIP
270
aquaporin in maize roots, using reverse genetic approaches. Overexpression of PIP2;5 under
271
the control of the 35S promoter led to a less than two-fold increase in the PIP2;5 protein level
272
in roots, where PIP2;5 is already highly expressed, and an approximately 10-fold increase in
273
leaves, where PIP2;5 is lowly expressed. This difference in PIP2;5 protein abundance
274
according to the organ suggests the existence of post-transcriptional or post-translational
275
regulation mechanisms that prevent an excess of PIP2;5 proteins according to the cell type.
276
Different cellular mechanisms modifying PIP abundance in the plasma membrane have been
277
reported (Chaumont and Tyerman, 2014; Maurel et al., 2015), and involve internalization of
278
PIPs from the plasma membrane for their degradation and/or recycling. Negative feedback of
279
PIP gene transcription could not be excluded either, even though it was not detected by
280
RT-qPCR. The decrease in PIP2;5 gene expression was obtained using the UniformMu
281
transposon mutated line UFMu00767 (McCarty et al., 2005; McCarty et al., 2013; Hunter et
282
al., 2014). In this line, the transposon was inserted in the second intron leading to an
283
important decrease in mRNA and protein levels. However, a very weak signal for PIP2;5
284
protein was still observed in roots, indicating that the line was not a complete knockout as
285
demonstrated by RT-qPCR using primers flanking the Mu insertion site allowing the
286
detection of a weak signal in pip2;5 KO samples (Supplemental Fig. S2).
287
The cortex cell Lpc in intact roots was dependent on PIP2;5 expression levels, with higher
288
and lower values in OE and KO lines, respectively. As no change in the abundance of other
289
PIP aquaporins was detected, this is a direct evidence that PIP2;5 facilitated the water
290
diffusion through the cell membranes. We previously showed a correlation between PIP
291
expression and the Lpc in roots. The higher abundance of PIPs, including PIP2;5, during the
292
day than during the night, or after a short (8 h) PEG treatment is correlated with variation in
293
the Lpc values (Hachez et al., 2012). Overexpression or knocking out PIP genes in other plant
294
species also results in higher or lower Lpc. For instance, in Arabidopsis thaliana pip2;2 KO
295
lines, Lpc of the root cortex cells is reduced by ~25% when compared with WT plants (Javot et
296
al., 2003). In contrast, Lpc is higher in PIP2;5 overexpressing Arabidopsis lines than in WT
297
under low temperature (Lee et al., 2012).
298
While the Lpr of pip2;5 KO plants was significantly decreased, no increase in Lpr was
299
recorded in PIP2;5 OE plants under well-watered conditions, indicating that the abundance of
300
PIP2;5 in the root cell membranes is not always correlated to the Lpr. The uncorrelated data
301
between PIP abundance, Lpc, and Lpr was previously observed in maize plants subjected to a
302
short PEG stress, which induces a higher PIP expression and a higher Lpc, but no change in Lpr
303
(Hachez et al., 2012). The composite water transport model (Steudle and Peterson, 1998)
304
assumes that Lpr is controlled by the hydraulic conductivity of apoplastic and cell-to-cell
305
pathways in parallel. We proposed that radial variations along each pathway critically affect
306
water transport due to the presence of gatekeeper cells at the beginning (epidermis and
307
exodermis) or the end (endodermis) of the radial path of water (Hachez et al., 2012;
308
Chaumont and Tyerman, 2014). While the current cell pressure probe technology did not
309
allow to directly verify this hypothesis, the use of the quantitative modeling framework
310
MECHA (Couvreur et al., 2018) gave us the opportunity to get a better understanding of the
311
mechanisms involved. The statistical comparison of our measured and simulated Lpr results
312
largely supports the hypothesis that plasma membrane permeability is not radially uniform in
313
WT (hypothesis A rejected, p < 0.01, Fig. 4C) but may be saturated in the endodermis and
314
exodermis (hypothesis B, Fig. 4E), thus following the observed radial aquaporin expression
315
patterns observed by Hachez et al. (2006). This is an important result, as none of the simplest
316
to most sophisticated models of radial water flow account for such heterogeneity, which does
317
affect Lpr in WT. On the other hand, knocking-out PIP2;5 gene expression led to a decrease in
318
Lpr suggesting that PIP2;5 is an essential actor facilitating radial water flow and controlling
319
whole root conductivity, as also observed in our simulations. Quantification of the level of
320
active PIP2;5 proteins in the endodermis would be very informative. Indeed, PIP2;5 gene
321
overexpression in this specific cell type would not lead to an increase in PIP2;5 protein due to
322
the above-mentioned post-translational mechanisms and that a maximum of active PIP2;5
323
(and other PIPs) was already reached. These predictions could be refined by generating maize
324
lines with deregulated PIP expression exclusively in the endodermis or the exodermis.
325
Maize lines expressing PIP2;5 cDNA under the control of p35S promoter showed a much
326
higher increase in PIP2;5 protein abundance in leaves than in roots. Considering that general
327
PIP expression in maize leaves is lower than in roots and that PIP2;5 is very lowly expressed
328
in leaves (Hachez et al., 2006; Hachez et al., 2008; Heinen et al., 2009), we hypothesize that
329
overexpressing PIP2;5 in this organ was not limited by similar post-transcriptional or
330
post-translational mechanisms observed in roots. Similar to what was observed in root cells,
331
an increase in Pos was measured for the leaf mesophyll cells overexpressing PIP2;5. We
332
previously demonstrated that transient expression of PIP2;5 in mesophyll cells increases the
333
water membrane permeability (Besserer et al., 2012). But in contrast to the effect of
334
PIP2;5OE on the whole root conductance, overexpression of PIP2;5 led to a higher Kleaf
335
compared to the Kleaf of WT-B104 plants (Fig. 3, D and E). A similar result was found, for
336
instance, in tomato plants overexpressing NtAQP1 (Sade et al., 2010). Inversely,
337
downregulation of PIP1 gene expression in Arabidopsis plants results in a decrease in the
338
mesophyll cell Pos and the Kleaf (Sade et al., 2014). Kleaf and the leaf radial water flow are
339
thought to be mainly controlled by the vascular bundle sheath cells surrounding the veins
340
(Sack and Holbrook, 2006; Shatil‐Cohen et al., 2011; Buckley, 2015). These cells can have
341
suberized walls, and are associated with a very low apoplastic flow. Arabidopsis plants, in
342
which PIP1 genes are specifically silenced in these cells, also exhibit decreased mesophyll
343
and bundle sheath Pos and decreased Kleaf (Sade et al., 2014). In our work, PIP2;5 OE plants
344
exhibited higher Pos of the mesophyll cells and possibly also of the bundle sheath cells,
345
resulting in an enhancement of conductance in bundle sheath, mesophyll cells, and Kleaf. The
346
observation that no difference in Pos and Kleaf was observed in pip2;5 KO plants was expected
347
since PIP2;5 is lowly expressed in leaf tissues (Hachez et al., 2008). Extending the MECHA
348
model to leaf tissues and cells will be very useful to address these questions related to leaf
349
water relations.
350
LER is controlled by plant hydraulic properties, including leaf and xylem water potential,
351
transpiration and root hydraulic conductivity (Caldeira et al., 2014), and involved hormonal
352
regulation (Nelissen et al., 2012; Avramova et al., 2015; Nelissen et al., 2018). Actually, we
353
considered that short-term variation of LER is a proxy of the change of xylem water potential
354
(Parent et al; 2009; Caldeira et al., 2014a). In a mild water deficit, a faster recovery of LER
355
after the early morning drop was recorded in PIP2;5 OE-4 compared with WT-B104 plants,
356
and a higher LER during the day and night was recorded. This LER response in PIP2;5 OE
357
plants can be correlated with a higher Kleaf and is consistent with the effect of Kleaf on LER
358
recovery observed in transgenic plants that differ in ABA concentration in the xylem sap,
359
showing differences in Lpr (Parent et al., 2009). Interestingly, while WT-B104 and PIP2;5 OE
360
plants had a similar Lpr in well-watered conditions, an osmotic stress led to an increased Lpr in
361
the OE plants, suggesting that changes in hydraulic conductance (Lpr and Kleaf) by PIP2;5
362
deregulation at the cell level translated into an overall change in whole plant hydraulic fluxes
363
(and leaf water potential) that affects the LER in mild-stress conditions. The observation that
364
the LER was not affected in well-watered plants implies that molecular mechanisms
365
controlling the overall root hydraulic conductance occur in some cell types resulting in a
366
non-uniform distribution of water permeability (see above) or at specific positions along the
367
root axis (e.g. the connection between root and leaf xylems).
368
In conclusion, deregulation of PIP2;5 aquaporin expression in maize plants highlighted
369
their role in controlling the root and leaf cell water permeability. However, understanding the
370
results obtained at the root level has required a hydraulic model developed at the cell and
371
tissue scales. MECHA allowed us to discard the hypothesis of radially uniform Lpc, and
372
suggests that, in well-watered conditions, the gatekeeper cells of WT plants have a saturated
373
membrane permeability. Transposing the model to the aerial part will definitely offer
374
possibilities to better understand the hydraulic properties of tissues and cells in diverse
375
conditions and investigate the role and regulation of aquaporins in specific hydraulic
376
processes. Indeed, we showed that the increase in the hydraulic conductance by
377
overexpressing aquaporin positively affects the LER under moderate stress conditions.
378 379
380 381
Materials and Methods
382
383
Plasmids and genetic construction to generate PIP2;5 overexpressing lines
384
The cauliflower mosaic virus p35S promoter was PCR-amplified from the pMDC43 vector
385
(Curtis and Grossniklaus, 2003) and cloned into the Gateway entry vector pDONR P4-P1R
386
(Invitrogen, Carlsbad, US) following the manufacturer’s instructions. The YFP cDNA
387
followed by the tnos terminator sequence was amplified together from the pH35GY vector
388
(Kubo et al., 2005) and cloned into the pDONR221 vector. Finally, a uracil-excision based
389
cloning cassette (USER) was added downstream of the p35S promoter sequence and
390
subsequently cloned into pDONR P2R-P3 (Nour-Eldin et al., 2006; Hebelstrup et al., 2010).
391
These three entry vectors were verified by sequencing and their inserts brought together by
392
LR recombination into the pBb7m34GW backbone vector suitable for maize (Zea mays)
393
transformation (Karimi et al., 2013) (https://gateway.psb.ugent.be). The latter contains the bar
394
selectable gene under the control of the p35S promoter that induces resistance to the herbicide
395
bialaphos and allows selecting transformed calli and shoots using phosphinotricin-containing
396
media. The cDNA encoding ZmPIP2;5 was PCR-amplified using USER primers (forward
397
primer: 5' GGTCTTAAUATGGCCAAGGACATCGAGG 3'; reverse primer: 5'
398
GGCATTAAUCTAGCGGCTGAAGGAGGCA 3') and inserted into the destination vector to
399
obtain the final vector. This plasmid was used to transform the hypervirulent EHA101
400
Agrobacterium tumefaciens strain (Hood et al., 1986) by heat shock and subsequently used for
401
transformation of maize immature embryos from B104 inbred line according to the method
402
described by Coussens et al. (2012), in collaboration with the Maize Transformation platform
403
of the Center Plant Systems Biology (VIB-Ghent University, Belgium). Briefly, immature
404
embryos were isolated from the ears 12 to 14 days after fertilization and co-cultivated with
405
EHA101 A. tumefaciens strain carrying the plasmid of interest for three days in the dark at
406
21℃. After co-cultivation, immature embryos were cultivated for one week without selection,
407
followed by a four-month period of subculturing on selective media containing increasing
408
amounts of phosphinotricin (starting from 1.5 mg/l to 6 mg/l) to select transformed
409
embryogenic calli in dark conditions at 25℃. After selection, the embryogenic calli were
410
transferred to bigger containers and placed in a growth chamber (24℃, 55μE.m-2.s-1 light
411
intensity, 16h/8h day/night regime) for transgenic T0 shoot/plantlet formation in vitro. Eleven
412
rooted transgenic T0 plants selected from independently transformed immature embryos
413
tested positive for the phosphinotricin acetyltransferase (PAT) enzyme (encoded by the bar
414
selectable marker gene) using the TraitChek Crop and Grain Test Kit (Strategic Diagnostics,
415
Newark, DE, USA. The presence of the transgenic ZmPIP2;5 cDNA insertion was also
416
confirmed by PCR amplification of genomic DNA using p35S forward primer (5'
417
CCACTATCCTTCGCAAGACCC 3') and T35S reverse primer (5'
418
GGTGATTTTTGCGGACTCTAGCAT 3'). The transgenic T0 plants were transferred to soil
419
and kept one month in a growth chamber (16h/8h day/night regime at 24 ℃, 55 µmol.m-2.s-1
420
light intensity, and 55% relative humidity) in 1 L pots for acclimation, and then moved to a
421
greenhouse (26 ℃/22 ℃, day/night temperature, 300 µmol.m-2.s-1 light intensity under
422
16h/8h day/night regime) in 10 L pots until flowering. A backcross with the B104 genotype
423
was performed by either pollinating ears of T0 plants with B104 pollen, or pollinating B104
424
ears with T0 plants pollen. The ears were then harvested 4 weeks after fertilization and dried
425
for several weeks at 25℃ before sowing the segregating T1 generation. The T2 generation
426
was generated by backcross between B104 and the heterozygous T1 plant in the greenhouse.
427
T1 and T2 generation heterozygous plants segregating for the ZmPIP2;5 transgene (PIP2;5
428
OE) and non-transgenic siblings (WT-B104) were used in this study. Finally, two independent
429
overexpressing lines (PIP2;5 OE-4 and PIP2;5 OE-13) with high ZmPIP2;5 protein content in
430
Western blotting analysis were used for all further measurements.
431 432
pip2;5 KO line
433
The pip2;5 KO line (UFMu00767, generated from the W22 inbred line, was found in
434
MaizeGDB (https://maizegdb.org/) and obtained from Uniform Mu stocks in “Maize Genetic
435
COOP Center” (http://maizecoop.cropsci.uiuc.edu/). To confirm the Mu transposon insertion
436
in PIP2;5 gene, gDNA was extracted from the second leaf of one-week old maize seedling,
437
with the extraction buffer TPS (100 mM Tris-HCl, pH 8.0, 10 mM EDTA, 1 M KCl). PCR
438
was performed using a Mu-TIR specific forward primer (McCarty et al., 2013) (Mu-Terminal
439
Inverted Repeat, TIR6: 5` AGAGAAGCCAACGCCAWCGCCTCYATTTCGTC 3`) and
440
PIP2;5 specific reverse primer (5` CGTCTACACCGTCTTCTCCG 3`) to detect the Mu
441
insertion, and with PIP2;5 specific primers (Forward: 5` AGGCAGACGATCCCAGCTT 3`,
442
Reverse: 5` CGTCTACACCGTCTTCTCCG 3`) to detect the PIP2;5 gene. Heterozygous
443
plants were self-pollinated and a segregation ratio of 1:2:1 for WT-W22, heterozygous, and
444
homozygous seeds was obtained. The WT-W22 and homozygous plants were used for the
445
measurements.
446 447
Growth conditions
448
Hydroponic culturing was conducted to obtain plant material for the measurements of the cell
449
hydraulic conductivity, root hydraulic conductivity, and leaf hydraulic conductance. Maize
450
seeds were surface sterilized using 2% (w/v) NaClO solution for 5 min, rinsed with distilled
451
water, and placed between two wet tissue papers in a square Petri dish (Greiner Bio-One,
452
Vilvoorde, Belgium). The seeds were put in the dark at 30℃ for 72 h. After germination, the
453
seedlings were transferred to a 2 L beaker with 1/2 strength nutrient solution (1.43 mM
454
Ca(NO3)2.4H2O, 0.32 mM KH2PO4, 0.35 mM K2SO4, 1.65 mM MgSO4.7H2O, 9.1 μM
455
MnCl2.4H2O, 0.52 μM (NH4)6Mo7O24.4H2O, 18.5 μM H3BO3, 0.15 μM ZnSO4.7H2O, 0.16
456
μM CuSO4.5H2O, and 35.8 μM Fe-EDTA. The nutrient solution pH was adjusted to 5.5 and
457
replaced every two days. The nutrient solution was aerated with the aid of aquarium diffusers.
458
After one week, the full-strength nutrient solution was used until the end of experiments. The
459
plants were grown in a growth chamber at a 16h/8 h light/dark cycle (25/18℃) and a daytime
460
light intensity of 200 μmol.m-2.s-1 at the top of the leaf level.
461
Soil culturing was conducted in the growth chamber or in the greenhouse. Maize seeds
462
were surface sterilized using 2% (w/v) NaClO solution for 5 min, rinsed with distilled water,
463
and placed in Jiffypots® (6 cm diameter), filled with 80% potting soil (DCM, Grobbendonk,
464
Belgium) and 20% vermiculite (Agra-vermiculite, Pull Rhenen, Netherland). After two weeks,
465
the seedlings were transferred to a 2 L pots (MCl 17, Pöppelmann, Geluwe, Belgium) filled
466
with the same substrate and vermiculite. After another 4 weeks, the plants were transferred to
467
a 10 L pot (MCl 29).
468 469
Protein extraction and Western blot
470
Microsomal membrane fractions were prepared as described by Hachez et al. (2006) with a
471
few modifications. Briefly, leaf (newly expanded leaf) or root (primary root) tissues from
472
one-week old maize seedling were flash-frozen in liquid nitrogen in aluminum foil and
473
grinded with mortar and pestle using 2 ml of extraction buffer (250 mM sorbitol, 50 mM
474
Tris-HCl pH 8, 2 mM EDTA,) containing freshly added 0.6% (w/v) polyvinyl pyrrolidone
475
K30, 1 mM phenylmethanesulfonate, 0.5 mM dithiothreitol, and supplemented with 2 μg.ml-1
476
of protease inhibitors (leupeptin, aprotinin, antipain, pepstatin, and chymostatin). Debris were
477
removed with a first centrifugation at 770g at 4℃ for 10 min and the subsequent supernatant
478
was centrifuged at 10,000g at 4℃ for 10 min. The supernatant was then centrifuged at
479
54,000g at 4℃ for 30 min and the resulting pellet (microsomal fraction) was resuspended in
480
50 μl of suspension buffer (330 mM sucrose, 5 mM KH2PO4, 3 mM KCl, pH 7.8) and
481
sonicated twice for 5 s. Total protein concentration was determined by the Bradford protein
482
assay (Bradford, 1976).
483
Twenty μg (for root) and 30 μg (for leaf) of total proteins were mixed with 6X Laemmli
484
buffer (240 mM Tris-HCl, pH 6.8, 6% w/v SDS, 30% w/v glycerol, 0.05% w/v bromophenol
485
blue) and freshly added 10% w/v dithiothreitol, in a total volume of 45 μl. They were
486
solubilized for 10 min at 60℃ were loaded in 12% acrylamide gels (Eurogentec, Seraing,
487
Belgium) for separation by electrophoresis (120V, ~1h). After transfer to a polyvinylidene
488
fluoride membrane (Trans-Blot® Turbo™ Mini PVDF Transfer Packs, Biorad, CA, USA), the
489
proteins were immunodetected using antisera raised against the amino-terminal peptides of
490
ZmPIP1;2, ZmPIP2;1/2;2, and ZmPIP2;5 (Hachez et al., 2006; Hachez et al., 2012). The
491
antibody raised against H+-ATPase (PMA) (Morsomme et al., 1996) was used as control to
492
normalize the protein level. The blotting signal was detected using an Amersham imager 600
493
(GE Healthcare, Chicago, USA). The signal was quantified with ImageJ software (National
494
Institutes of Health, http://rsb.info.nih.gov/ij) and normalized with the signal of PMA.
495 496
RNA extraction and reverse transcription quantitative PCR (RT-qPCR)
497
One-week old maize seedlings were used for RNA extraction and RT-qPCR. The whole newly
498
expanded leaf and primary root were harvested and immediately frozen in liquid nitrogen, and
499
the samples were stored at -80℃ until RNA extraction. RNA extraction was performed with
500
the Spectrum™ Plant Total RNA Kit (Sigma-Aldrich, Mo, US) according to the
501
manufacturer’s instructions. DNAse I (Sigma-Aldrich) digestion was performed directly on
502
the column during RNA extraction according to the manufacturer’s recommendations. The
503
RNA concentrations and the quality of each sample were measured with a Nanodrop
504
ND-1000 (Isogen Life Science, Utrecht, Netherland), and 1.5 μg of total RNA was used for
505
reverse transcription and the cDNA synthesis was performed with the Moloney Murine
506
Leukemia Virus Reverse Transcriptase (M-MLV RT) kit (Promega, Leiden, Netherland)
507
according to the manufacturer’s instructions. RT-qPCR was performed in 96-well plates using
508
a StepOnePlus™ Real-Time PCR System (Life Technologies, Foster City, CA, USA) in a
509
volume of 20 μl containing 10 μl of RT-qPCR Mastermix Plus for SYBR Green I (Eurogentec,
510
Liège, Belgium), 1 μl of forward and reverse primers (Supplemental Table S1), 1 μl cDNA,
511
and 7 μl DEPC-H2O. The PCR cycle program was 2 min at 50℃, 10 min at 95℃ for DNA
512
polymerase activation, and 40 cycles of 15 s at 95℃ and 60 s at 60℃. The 2-ΔCt method was
513
used to analyze the relative expression of 6 ZmPIP1s (ZmPIP1;1, ZmPIP1;2, ZmPIP1;3,
514
ZmPIP1;4, ZmPIP1;5, and ZmPIP1;6) and 6 ZmPIP2s (ZmPIP2;1, ZmPIP2;2, ZmPIP2;3,
515
ZmPIP2;4, ZmPIP2;5, and ZmPIP2;6). Three reference genes, ACT1, EF1-α, and
516
polyubiquitin were used to normalize the expression.
517 518
Cell pressure probe measurement
519
One-week old maize seedlings were used for cell pressure probe measurements, according to
520
Hachez et al. (2012) and Volkov et al. (2006). Briefly, the plant and primary root was
521
maintained in a Petri dish containing the nutrient solution and the root region near to the last
522
lateral root (5-7 cm near the root tip) was measured. Cell turgor pressure (P), T1/2 (half-time of
523
water across cell membrane), and pressure change value (ΔP) were recorded. These
524
parameters for three to five cells from each plant were measured and three to five plants were
525
analyzed. After the measurements, average values of cell volume and cell surface area were
526
calculated through microscopic analyses of 7-30 cells from the 3rd to 6th cell layer taken at 5-7
527
cm from the root tip. Cell osmotic pressure was checked with micro-osmometer (Advanced,
528
model 3300, Norwood, Massachusetts, USA). In this study, a cell osmotic pressure of 0.88
529
MPa was used for all calculations. Cell elastic modulus (ε) and cell hydraulic conductivity
530
(Lpc) were calculated according to Volkov et al. (2006).
531 532
Root hydraulic conductivity (Lpr) and leaf hydraulic conductance (Kleaf) measurement
533
Three-week old maize plants growing in phytotron were used for root hydraulic conductivity
534
(Lpr) measurements, according to Ding et al. (2015). In the phytotron, the light cycling was
535
from 06:00 to 22:00. In the morning between 09:00~11:00 (measurements beginning three
536
hours after light onset), the primary root was cut and connected to a hydraulic conductance
537
flow meter (HCFM, Decagon Devices, Pillman, WA, USA). The connection was performed as
538
quickly as possible (< 1 or 2 min) to minimize the effect of the root excision on the Lpr as
539
previously reported by Vandeleur et al. (2014). Transient method was used to recording the
540
value change of flow rate (F) and applied pressure (Pi), and the slope rate (Kr, Kg.s-1.MPa-1)
541
was analyzed between the correlation of F and Pi. For one root, three to five replications were
542
performed for the measurement. After the measurement, the primary root was scanned with
543
Epson perfection V33 scanner (EPSON, Japan) and the root surface area was analyzed with
544
ImageJ (National Institutes of Health, http://rsb.info.nih.gov/ij). Then the Kr was normalized
545
with the root surface area for the Lpr calculation. For PEG treated plants, 10% (w/v)
546
PEG-6000 was added to the nutrient solution in hydroponic culture, and Lpr was measured
547
after 2- and 4-h.
548
Leaf hydraulic conductance (Kleaf) was measured between 13:00~15:00 (measurements
549
beginning seven hours after light onset) with HCFM (Tyree et al., 2004; Ferrio et al., 2012)
550
using three-week old maize plants. The newly expanded leaf was cut with the leaf sheath, and
551
coated surrounding a plastic stick covered by UHU®patafix (UHU, Baden, Germany). Then a
552
tape of polytetrafluoroethylene film was wrapped around the leaf sheath. After, the leaf sheath
553
was excised under water and connected with the HCFM. Approximately 0.2 MPa pressure
554
was applied in the system and quasi-steady state method was used to record the flow rate and
555
conductance by every 8 s during 10-30min until the conductance was constant. During the
556
measurement, the leaf was immersed in water to stop transpiration. After the measurement,
557
the leaf was scanned with Epson perfection V33 scanner (EPSON, Japan) and leaf area was
558
analyzed with ImageJ software. The hydraulic conductance was normalized with the leaf area
559
to calculate the Kleaf.
560 561
Protoplast swelling assay
562
Leaf mesophyll protoplasts were isolated from newly expanded leaves of three-week old
563
maize plants, and swelling assay was performed according to Moshelion et al. (2004) and
564
Shatil-Cohen et al. (2014). The leaf abaxial side was scratched with a glass-paper, and then
565
the leaf was cut into small sections and transferred to the digestion buffer (the scratched side
566
contact with the buffer), including 0.6% (w/v) cellulose R10 (Duchefa Biochemine, Haarlem,
567
Netherland), 0.1% (w/v) pectolyase (Sigma-Aldrich, Mo, US), 0.3% (w/v) Macerozyme R10
568
(Duchefa Biochemine, Haarlem, NL), 5 mg.ml-1 bovine serum albumin, and 5 mg.ml-1
569
polyvinyl pyrrolidone K30. The analysis of the osmotic water permeability coefficient was
570
according to Shatil-Cohen et al. (2014).
571 572
Leaf elongation rate measurement
573
Leaf growth was measured in the high throughput phenotyping platform Phenodyn (Sadok et
574
al., 2007) of the LEPSE laboratory in Montpellier
575
(https://www6.montpellier.inra.fr/lepse/M3P). Plants were grown in one PVC column, filled
576
with clay balls, which allows to tare columns at 1.2 kg. Then, columns were filled with 4.4 kg
577
of a mix of loam (5-10%) and soil. Plants were daily watered with nutritive solution.
578
Sampling of the newest leaf was carried out at 4-leaf stage and a PCR was made in order to
579
characterize the transgenic PIP2;5 OE plants. After the characterization, three plants were
580
kept in the column, including either three WT-B104 or PIP2;5 OE plants or a mix of them
581
(either two WT-B104 or two PIP2;5 OE) for the LER measurement. Plants were grown under
582
a 14 h light / 10 h dark cycle at 19-26℃ (day, mini-maxi)/19-21℃ (night, mini-maxi) in
583
greenhouse. Well-watered conditions were kept until plants reached the four leaf
584
developmental stage. Then a progressive water deficit was applied. Each pot was placed on a
585
scale with automated irrigation to impose the targeted soil water potential. LER was measured
586
when the tip of the 6th leaf appeared above the whorl and lasted until the appearance of leaf 8.
587
LER was expressed in thermal time, via equivalent days at 20℃ according to Parent et al.
588
(2010). Ψleaf was measured with a pressure chamber between 12:00-14:00 in the greenhouse
589
with non-expanding leaves.
590 591
Inference on radial patterns of cell membrane permeability
592
In order to test hypotheses on the radial pattern of plasma membrane permeability, we
593
simulated how measured cell-scale permeability (Lpc) translates into root hydraulic
594
conductivity (Lpr) for each pattern, and compared the distributions of simulated and measured
595
Lpr.
596
Assuming that the axial resistance to water flow is negligible, the simulation framework
597
MECHA (Couvreur et al., 2018) estimates Lpr from root transverse anatomy and subcellular
598
scale hydraulic properties. In order to plug the measured Lpc into the model, we partitioned it
599
into its two main components: the plasma membrane hydraulic conductivity (kAQP, including
600
the contribution of aquaporins) and the conductance of plasmodesmata per unit membrane
601
surface (kPD). The latter parameter was assumed to equal 2.4×10-7 m.s-1.MPa-1 following
602
